TL1-A can engage death receptor-3 and activate NF-kappa B in endothelial cells
© Wang et al.; licensee BioMed Central Ltd. 2014
Received: 4 April 2014
Accepted: 31 October 2014
Published: 16 November 2014
Death receptors (DRs) play an important role in renal pathology. We have shown that DR3 is inducibly expressed on renal tubular epithelial cells in the setting of inflammatory injuries. In this study we investigate the expression of DR3 in renal endothelial cells and their response to TL1A, the only known ligand of DR3.
We did RT-PCR, flow cytometry and subcellular immunoblotting to examine the expression and function of DR3 in cells in vitro. We did organ culture of human and mouse tissue to examine expression and signal of DR3 in vivo.
DR3 is expressed in some interstitial vascular endothelial cells (EC) in human kidney in situ; these EC also respond to its ligand TL1A by activating NF-κB. Very low levels of DR3 can be detected on the cell surface of cultured human umbilical vein (HUV) EC, which do not respond to TL1A. HUVEC transfected to overexpress DR3 become responsive to TL1A, assessed by IκBα degradation and E-selectin induction, indicating that the signaling components needed for DR3 responsiveness are expressed. TL1A induces NF-κB activation in EC in renal and cardiac tissue from wild type but not DR3 knock-out mice.
TL1A and DR3 activate NF-κB in vascular endothelial cells, and can be an important regulator of renal interstitial vascular injury.
KeywordsKidney Endothelial cells Death receptor Inflammation
Death receptors (DRs) of the tumor necrosis factor receptor (TNFR) superfamily include TNFR1, Fas, DR3, DR4, DR5 and DR6. The members of DR family each contain an intracellular death domain (DD) that binds to adaptor proteins which also contain DDs and initiate apoptotic cell death . The DD of DR3 shares the highest homology with that of TNFR1 (47%) , and, like TNFR1, interacts with the DD-containing silencer of death domain (SODD) in the absence of ligand, whereas the other four DRs do not [3, 4]. Furthermore, upon ligand binding, only DR3 and TNFR1 recruit the adaptor TNFR-associated death domain (TRADD) protein and assemble a TRADD-dependent signalling complex that rapidly triggers NF-κB and c-Jun N-terminal kinase (JNK) activation; after a delay of several hours the TRADD signalosome recruits Fas-associated DD (FADD) protein that may trigger apoptosis . However, in contrast to TNFR1, which is expressed on a wide range of cell types, DR3 expression is more restricted  and most commonly associated with T cells. Two different species of messenger RNA (mRNA) encoding a transmembrane form of DR3, DR3 and DR3β have been isolated from of a panel of human lymphoid cell lines. Eleven splice variants of DR3 mRNA have been identified in peripheral blood lymphocytes, and upon activation by phytohaemagglutinin (PHA), the transcript level is up-regulated [7, 8]. Much less is known about regulation and expression of DR3 protein or about its isoforms in non-lymphoid cells. We have previously reported that DR3 mRNA and protein are induced on non-lymphoid cells of human kidney allografts undergoing acute cellular rejection . Specifically, we observed that DR3 is predominatly up-regulated in tubular epithelial cells but we also observed expression of DR3 in some peritubular endothelial cells (EC), albeit not in glomerular EC.
The only known ligand for DR3 is TNF-like molecule 1A (TL1A), a TNF superfamily member, and mRNA encoding this protein is most abundantly expressed in kidney tissue; it is a long variant of TL1 (also called vascular endothelial growth inhibitor, VEGI) . Both TL1A mRNA and protein are also up-regulated by the same pathological processes that control DR3 expression in renal epithelial cells . In situ hybridization suggested that EC are a major source of TL1A in the human kidney. The TL1A-DR3 system may thus play an important role in the vasculature in renal injury and inflammation. Human umbilical vein (HUV) ECs are a major source of TL1A synthesis among cultured cell types, but it is not known whether EC respond to this ligand. We have investigated fully the expression and function of DR3 in human EC in this study.
Mouse monoclonal anti-human DR3 was purchased from Chemicon (Southampton, U.K). Rabbit anti-human IκBα and goat anti-human DR3 antibodies were from Santa Cruz (Wiltshire, U.K). Horse anti-goat and goat anti-rabbit horseradish peroxidase (HRP)-conjugated antibodies and Vectashield Mounting Media were from Vector Laboratories Ltd (Peterborough, U.K). Rabbit anti-human NF-κB p65 was purchased from Serotec (Oxford, U.K). Rabbit anti-human phospho-NF-κB p65 was from New England Biolab (Hertfordshire, U.K). Proteinase inhibitor cocktail was purchased from Roche Diagnostics Ltd (East Sussex, U.K). Human recombinant TL1A, human recombinant TNF-α and goat anti-human DR3 antibody were purchased from R&D Systems Europe (Abingdon, U.K). Chicken anti-rabbit Alexafluor488, goat anti-mouse Alexafluor568, Lipofectin reagent and Opti-MEM media were from Invitrogen Ltd (Paisley, U.K). The ECL system was from Amersham Pharmacia Biotech UK Ltd (Bukinghamshire, U.K).
Unless otherwise indicated, all reagents were from Sigma-Aldrich Company Ltd (Dorset, U.K).
All experiments using human tissue were performed with written, informed consent of patients and the approval of Local Ethical Committee which is East of England - Cambridge Central Research Ethics Committee and Addenbrooke’s Hospital Tissue Bank. Renal tissues was obtained from the uninvolved pole of kidney excised less than half an hour before because of renal tumours or from time zero biopsy of kidney transplant.
Kidney organ culture
Duplicate 1 mm3 fragments of kidney tissue were placed in flat-bottomed 96-well tissue culture plates and immediately immersed in medium M199 containing 10% heat inactivated bovine fetal calf serum (FCS) and 2 mM L-glutamine. Tissue was incubated for 3 hours at 37°C with either culture media alone or with 5 ng/ml TNF-α (TNF) or 0.2 μg/ml TL1A. Half of the harvested tissue was cryoprotected in 30% sucrose in 0.1 M phosphate buffer and snap frozen in isopentane-cooled liquid nitrogen and half was immersed in 4% paraformaldehyde in 0.1 M PIPES buffer pH 7.6 for 1.5 hours at 4°C and processed for paraffin-wax embedding.
Immunolabelling of tissue
8 μm-thick cryosections of kidney tissue processed as above were permeabilized in cold methanol at -20°C for 5 minutes, washed in Milli-Q water and rinsed in 0.1 M Tris–HCl buffer pH 7.5 containing 0.01% TWEEN-20 (TBS) prior to incubation with blocking buffer (10% FCS in TBS) for 10 minutes. Sections were immunolabelled as previously described  Sections were then washed and mounted in Vectashield Mounting Media and imaged with Leica SPE confocal laser scanning microscope (Leica Microsystem Ltd, Milton Keynes, U.K).
Human umbilical vein EC (HUVEC) were from Lonza (Cambrdige, UK) and serially cultured as previously described . Human dermal microvascular cells (HDMEC) and human pulmonary artery endothelial cells (HPAEC) obtained from Life Technology (Paisley, UK) were cultured following the supplier’s instruction. Cells were used at passages 1–3. Such cultures are free of detectable leukocytes by immunostaining for CD45. TF-1 human erythroleukemia cells obtained from American Tissue Culture Centre (LGC Promochem, Middlesex, U.K) were cultured in media as supplier required.
Total RNA was isolated from cells using RNeazy Mini Kit (QIAGEN Ltd, West Sussex, U.K). 1 μg total RNA was amplified with DR3 forward (5′-GTAGCCCCAGGTGTGACTGT-3′) and DR3 reverse (5′-GCTTGAGCATCTCGTACTGC-3′) using Access RT-PCR system (Promega, Southampton, U.K); the PCR product was visualized by 1% low melting temperature agarose gel in Tris-Acetate-EDTA buffer before purified with Wizard PCR Preps (Promega, U.K). The purified DNA was sequenced in Lark Technologies Inc (Essex, U.K).
Measurement of cell surface receptor expression by flow cytometry
HUVEC were seeded into 6-well tissue culture plate (1.5 × 105 cells per well), and 24 hours later the confluent cells were treated with IFN-γ 500 U/ml, or TNF-α 500U/ml or the combination of both for 24 and 48 hours. Cells were harvested using a non-enzymatic cell suspension solution. TF-1 cells (106/ml) were centrifuged down. Cells were immunolablled and Fixed before analyzed by flow cytometry using FACSCalibur machine (BD Biosciences, Oxford, U.K) as previously described . Data were analyzed using WinMDI 2.8 software.
HUVEC were washed once with PBS and scraped into homogenization buffer (10 mM Tris HCl, pH 7.4; 3% sucrose w/v; proteinase inhibitor cocktail, 1 mM PMSF). TF-1 cells were pelleted by centrifugation, washed once with PBS and resuspended in homogenization buffer. All steps were performed at 4°C as previously described . The protein concentration of each fraction was determined by BCA protein assay kit (Pierce, Chester, U.K). Samples were stored at -80°C before analysis by immunoblotting.
Generation of DR3-flag construct and transfection
The open reading frame of the human DR3 gene was isolated by reverse transcription using the forward primer 5′-TAGCGAATTCAATGGAGCAGCGGCCGCGGG-3′ and the reverse primer 5′-GCGCTCTAGAAACCGTACTTAGGGCTTCTGC-3′. The amplified product was ligated in-frame into the pFlag-CMV1 vector (Sigma-Aldrich Company Ltd, Dorset, U.K). DR3-Flag constructs were introduced into passage 1 HUVEC by transient transfection. In brief, HUVEC grown to 70% confluence on 100 mm diameter plastic culture plates or 6-well plates were transfected approximately 2 hours with DR3-Flag using Lipofectin reagent in Opti-MEM media following manufacture’s instruction. Cells were cultured for another 48 hours. Cells transfected with empty vector or DR3-Flag were treated with TNF 5 ng/ml or TL1A 0.2 μg/ml for 2 or 4 hours. Cells were then collected for analyzing by flow cytometry as described above.
HUVEC transfected with control vector or DR3-Flag were stimulated with 0, 0.02, 0.2, 2 μg/ml TL1A or 0, 1, 5, 10 ng/ml TNF for 30 minutes and then washed once with ice cold PBS before lysed in lysis buffer (62.5 mM Tris, 2% SDS, 10% glycerol, 10 mM sodium orthovanadate, 10 mM sodium fluoride, proteinase inhibitor cocktail). Samples (20 μg for IκBα, 50 μg for subcellular fraction) were immunoblotted and detected as previously described . Serial dilution of samples for immunoblotting confirmed that the density of bands was within the linear range of detection.
Mouse kidney and heart
Protocols involving animal were approved by the U.K home office and the Cambridge University Local Ethical Committee. C57Bl/6 DR3-/- mice were crossed once into a CD1 background, and the F1 heterozygote progeny were crossed to yield DR3+/+ wild type (WT) and DR3-/- knockout (KO) littermates. Animals were killed and the kidneys and hearts were harvested and processed as described in the previous section.
The significance of difference between experimental values was assessed by means of the paired Student’s t test.
Activation of NF-κB by TL1A in kidney organ culture
Expression of DR3 in cultured EC
We have found that the magnitude of IκBα degradation following TNF stimulation is related to the number of cell surface TNF receptors [12, 13]. We tried to up-regulate EC’s surface DR3 using cytokines, including TNF-α and IFN-γ alone and in combination without success. Then we transiently transfected HUVEC to increase expression of DR3, using a DR3-Flag construct, followed by flow cytometry and immunoblotting to confirm up-regulation of DR3. Although the major band expressed after transfection was a shorter form of DR3 (as recognized by both anti-DR3 antibody and anti-Flag anti-body), the full length form of DR3 did increase significantly after transfection (Figure 2E).
Function of DR3 in EC
To determine whether EC from other vascular beds constitutively signal through DR3, we studied DR3 response to treatment with TL1A in HDMEC and HPAEC. HDMEC and HPAEC degraded IκBα in response to treatment with TNF, but not TL1A. Consistent with this we could not detect full length DR3 by immunoblotting in HDMEC and HPAEC (see Additional file 1).
Responses of EC to TL1A in DR3 knockout mouse
In this study we have shown for the first time that human EC express full length DR3, which supports a more widespread cellular expression of DR3 than its original description as a lymphoid cell receptor [14–16]. Moreover, EC have all the machinery necessary to respond upon DR3 up-regulation. Exogenous TL1A can induce NF-κB activation in EC both in vitro and in vivo.
TNF superfamily members play an important role in renal injury , and the differential expression of TNF receptor (TNFR) superfamily members may have functional implications . TNFRs are widely expressed in glomerular and interstitial EC, while DR3 is detected only in some interstitial EC. Our results show that, like TF-1 cells, HUVEC express both DR3 transcript and protein. Although we were unable to identify a pathophysiological stimulus that would up-regulate full length DR3 in EC, we were able to increase surface expression of DR3 by transfection, which resulted in IκBα degradation by TL1A treatment. This potential function of DR3 in EC is supported by our in situ study of EC in TL1A-treated kidney tissue.
Pre-ligand-binding assembly of TNFR into trimer complexes is critical in TNF mediated signaling , which is based on a ligand-receptor trimerisation. It is possible that TL1A signaling does not occur in untreated HUVEC because there is both insufficient level of DR3 on the surface, and other forms of non-signaling DR3 interfere with TL1A binding. The lack of response to TL1A cannot be attributed to inability of DR3 to signal IκBα degradation in HUVEC, as expressing full length DR3 by transfection allowed TL1A to degrade IκBα. The lower molecular weight band detected by immunoblotting with anti-flag and anti-DR3 after transfection may be an isoform of DR3 that is able to bind TL1A and interfere with its signalling, as reported for other death receptors like DR4 and DR5 . This DR3 isoform differs from the reported soluble form of DR3  as it is detected only in the membrane fraction and not the soluble fraction. It is unlikely to be the signaling isoform inducing IκBα degradation as it differs from the isoform expressed in TF-1 cells. Moreover, the relatively abundant expression of this shorter isoform after transfection did not make HUVEC more responsive to TL1A than TF-1 cell, which in turn indicate that it is full-length DR3 that TL1A signals through. Further studies on isoforms of DR3 are needed to support our hypothesis. Our results are consistent with Migone et al., when they over expressed TL1A and DR3 in 293T cells, NF-κB activation could be detected, whereas VEGI failed to do so .
NF-κB factors belong to the Rel family of transcription factor; they mediated biological activity through classical and alternative pathways. Our results in tissue support DR3 signaling through the classical pathway similar to TNFR1, which induces IκBα degradation and RelA/p65 activation . Although we were unable to demonstrate activation of RelA/p65 in TF1 cells following IκBα degradation, we have shown that TL1A induces RelA/p65 activation in EC in situ. Furthermore, we demonstrated TL1A induced E-selectin expression in DR3 transfected HUVEC, indicating functional activation of NF-κB in these cells. We have previously observed differences in death receptor signaling in situ compared to in vitro . Whether the alternative pathway is involved requires further studies. Although TL1A-DR3 interaction has been extensively studied in autoimmune diseases such as inflammatory bowel diseases, experimental allergic encephalomyelitis and rheumatoid arthritis [21–23], its physiological role in immune reaction is still unclear. Furthermore it has been shown to have functions outside of the immune system, including atherosclerosis and aging [24, 25]. Both TL1A and DR3 expression levels are low under physiological conditions, but can be acutely up-regulated in inflammatory reaction. As EC also express TL1A, the dynamic balance of the ligand and receptor in endothelial systems could be critical for maintaining normal function of these cells. Over expression of either ligand or receptor could lead to inflammation or apoptosis . In normal kidney where most cells are negative for DR3, TL1A may be unable to induce signaling. However, in human allograft rejection infiltrating mononuclear cells, which express high levels of DR3, are able to respond to TL1A, the DR3-TL1A interaction may form a positive feedback loop inducing an inflammatory reaction.
EC express full length and functional DR3. The TL1A and DR3 system may play an important role in vascular injury in allograft rejection and atherosclerosis where significant increases in both TL1A and DR3 have been reported.
This work was supported by grants from the National Natural Science Foundation of China (30971120), British Heart Foundation, NIHR Cambridge Biomedical Research Centre, and the National Institutes of Health. We thank Yi Fan for technical help in DR3 vector preparation.
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